Ribosomes: Structure, Function, and Composition
Ribosomes are ribonucleoproteins which are present in both prokaryotes and eukaryotes. They consist of about two-thirds RNA and one-third protein. Ribosomes are essential for protein synthesis. In the last step of the gene expression pathway, ribosomes translate the genomic information encoded in a messenger RNA into protein (Garrett, et al., 2000).
Ribosomes are complexes of two nonequivalent ribonucleoprotein subunits. The larger subunit (“large ribosomal subunits”) is about twice the size of the smaller (“small ribosomal subunits”). The small ribosomal subunit binds mRNA and mediates the interactions between mRNA and tRNA anticodons on which the fidelity of translation depends. The large ribosomal subunit catalyzes peptide bond formation—the peptidyl-transferase reaction of protein synthesis—and includes two different tRNA sites: the A site for the incoming aminoacyl-tRNA, which is to contribute its amino acid to the growing peptide chain, and the P site for peptidyl-tRNA complex, i.e. the tRNA linked to all the amino acids that have so far been added to the peptide chain. The large ribosomal subunit also includes a binding site for G-protein factors that assist in the initiation, elongation, and termination phases of protein synthesis. The large and small ribosomal subunits behave independently during the initiation phase of protein synthesis; however, they assemble into complete ribosomes when elongation is about to begin.
The molecular weight of the prokaryotic ribosome is about 2.6×106. In prokaryotes, the small ribosomal subunit contains a 16S (Svedberg units) rRNA having a molecular weight of about 5.0×105. The large ribosomal subunit contains a 23S rRNA having a molecular weight of about 1.0×106 and a 5S rRNA having a molecular weight of about 4.0×105. The prokaryotic small subunit contains 21 different proteins and its large subunit, 31 proteins. The large and small ribosomal subunits together make a 70S ribosome in procaryotes.
Eukaryotic ribosomes are bigger than their prokaryotic counterparts. The large and small subunits together make an 80S eukaryotic ribosome. The small subunit of an eukaryotic ribosome includes a single 18S rRNA, while the large subunit includes a 5S rRNA, a 5.8S rRNA, and a 28S rRNA. The 5.8S rRNA is structurally related to the 5′ end of the prokaryotic 23S rRNA and the 28S rRNA is structurally related to the rest (Moore, 1998). Eukaryotic ribosomal proteins are qualitatively similar to the prokaryotic ribosomal proteins; however, the eukaryotic proteins are bigger and there are more of them (Moore, 1998).
Structural Conservation of the Large Ribosomal Subunit
While the chemical composition of large ribosomal subunits varies significantly from species to species, the sequences of their components provide unambiguous evidence that they are similar in three-dimensional structure, function in a similar manner, and are related evolutionarily. The evolutionary implications of the rRNA sequences data available is reviewed in the articles of Woese and others in part II of “Ribosomal RNA. Structure, Evolution, processing and Function in Protein Biosynthesis”, Zimmermann and Dahlberg, eds, CRC Press, Boca Raton, Fla., 1996. The article by Garret and Rodriguez-Fonseca in part IV of the same volume discusses the unusually high level of sequence conservation observed in the peptidyl transferase region of the large ribosomal subunit. Archeal species like H. marismortui have ribosomes that resemble those obtained from eubacterial species like E. coli in size and complexity. However, the proteins in their ribosomes are more closely related to the ribosomal proteins found in eukaryotes (Wool, I., Chan, Y.-L., & Gluck, A., Biochem. Cell Biol. 73, 933-947 (1995)).
Because of the high level of sequence conservation that characterizes the active site regions of ribosomes from different species, knowledge of the three-dimensional structure of a large ribosomal subunit from a single species belonging to a single kingdom, e.g. that of H. marismortui, will enable those skilled in the art both to understand the function of the critical regions of ribosomes from other species, regardless of kingdom. Thus it should be possible to for such an individual to produce useful models for the functionally significant regions of the ribosomes of higher organisms like humans and of the ribosomes from the bacteria that are their pathogens, and to understand how they might differ.
Determination of the Structure of Ribosomes
Much is what is known about ribosome structure derives from physical and chemical methods that produce relatively low-resolution information. Electron microscopy (EM) has contributed to the understanding of ribosome structure ever since the ribosome was discovered. In the 1970s, low resolution EM revealed the shape and quaternary organization of the ribosome. By the end of 1980s, the positions of the surface epitopes of all the proteins in the E. coli small subunit, as well as many in the large subunit, had been mapped using immunoelectron microscopy techniques (Oakes et al., 1986; Stoeffler et al., 1986). In the last few years, advances in single-particle cryo-EM and image reconstruction have led to three dimensional reconstructions of the E. coli 70S ribosome and its complexes with tRNAs and elongation factors at resolutions between 15 Å and 25 Å (Stark et al., 1995; Frank et al., 1995; Stark et al., 1997a; Agrawal et al., 1996; Stark et al., 1997b). Additionally, three-dimensional, electron microscopic images of the ribosome have been produced at resolutions sufficiently high so that many of the proteins and nucleic acids that assist in protein synthesis can be visualized bound to the ribosome (Agrawal et al., 2000), and earlier this year an approximate model of the RNA structure in the large subunit was constructed to fit a 7.5 Å resolution electron microscopic map of the 50S subunit from E. coli as well as biochemical data (Mueller et al., 2000).
While the insights provided by electron microscopy have been useful, it has long been recognized that a full understanding of ribosome structure would derive only from X-ray crystallography. Crystallization studies of the ribosome began two decades ago by Ada Yonath and coworkers opened the possibility of using X-ray crystallography to determine the structure of the ribosome at atomic resolution. This was a challenging enterprise. Crystals of ribosomes have been especially difficult to obtain because of their huge size and their lack of internal symmetry. Moreover, since their surface is composed of highly degradable RNA and loosely held proteins, ribosomes exhibit inherent flexibility and instability. In 1979, Yonath and Wittman obtained potentially useful crystals of ribosomes and ribosomal subunits (Yonath et al., 1980). Ribosomal crystals proved to be extremely sensitive to radiation even at cryo-temperature when using bright Synchroton X-ray beam required for high resolution data collection (Weinstein et al., 1999). They are also characterized by low level isomorphism, fluctuations in the unit cell dimensions, deformed spot-shape, and nonisotropic mosaicity (Weinstein et al., 1999). By the mid 1980s, Trakanov et al. (1987), a group in Puschino, were also preparing ribosome crystals for X-ray crystallography. Maskowski et al. (1987) were the first to obtain crystals of 50S ribosomal subunit from Haloarcula marismortui. In 1991, van Bohlen et al. (1991) reported an important improvement in the resolution of the diffraction data obtainable from the crystals of the 50S ribosomal subunit of H. marismortui. 
In 1995, Schlunzen et al. (1995) reported low resolution electron density maps for the large and small ribosomal subunits from halophilic and thermophilic sources. However, the low resolution electron density maps of Schlunzen et al. (1995) are incorrect (Ban et al., 1998).
The first electron density map of the ribosome that showed features recognizable as duplex RNA was a 9 Å resolution X-ray crystallographic map of the large subunit from Haloarcula marismortui published two years ago (Ban et al., 1998). A year later, extension of the phasing of that map to 5 Å resolution made it possible to locate several proteins and nucleic acid sequences the structures of which had been determined independently (Ban et al., 1999).
At about the same time, using similar crystallographic strategies, a 7.8 Å resolution map was generated of the entire T. thermophilus ribosome showing the positions of tRNA molecules bound to its A, P, and E (protein exit site) sites (Cate et al., 1999), and a 5.5 Å resolution map of the 30S subunit from Thermus thermophilus was obtained that allowed the fitting of solved protein structures and the interpretation of some of its RNA features (Clemons et al., 1999). Subsequently, an independently determined, 4.5 Å resolution map of the T. thermophilus 30S subunit was published, which was based in part on phases calculated from a model corresponding to 28% of the subunit mass that had been obtained using a 6 Å resolution experimental map (Tocilj et al., 1999). The subunit packing interpretation of the two 30S structures is not the same, even though the crystals used by the two groups appear to be identical.
The Need for Higher Resolution to Obtain the Atomic Structure for the 50S Ribosomal Subunit
Although the prior art provides crystals of the 50S ribosomal subunit and 9 Å and 5 Å resolution X-ray crystallographic maps of the structure of the 50S ribosome, the prior art crystals and X-ray diffraction data are not sufficient to establish the 3D structures of all 31 proteins and 3,043 nucleotides of 50S. Thus, the prior art crystals and maps are inadequate for the structure-based design of active agents, such as herbicides, drugs, insecticides, animal poisons, etc.
More detailed, higher resolution X-ray crystallographic maps are necessary in order to determine the location and 3D structure of the proteins and nucleotides in ribosomes and ribosomal subunits, particularly for the 50S ribosomal subunit. Such high resolution maps would enable the design of various useful active agents, such as herbicides, drugs, insecticides, animal poisons, etc.
For example, an accurate molecular structure of the 50S ribosomal subunit will not only enable further investigation and understanding of the mechanism of protein synthesis, but also the development of effective therapeutic agents and drugs that modulate (i.e., promote or inhibit) protein synthesis.
Location of the Peptidyl Transferase Site in the Large Ribosomal Subunit
It has been known for thirty-five years that the peptidyl transferase activity responsible for the peptide bond formation that occurs during messenger RNA-directed protein synthesis is intrinsic to the large ribosomal subunit (Traut et al., 1964; Rychlik, 1966; Monro, 1967; Maden et al., 1968) and it has been understood for even longer that the ribosome contains proteins as well as RNA. In bacteria, for example, the large ribosomal subunit contains ˜35 different proteins and two RNAs (Noller, 1984; Wittmann-Liebold et al., 1990). These findings posed three related questions. Which of the almost 40 macromolecular components of the large ribosomal subunit contribute to its peptidyl transferase site, where is that site located in the large subunit and how does it work?
By 1980, the list of components that might be part of the ribosome's peptidyl transferase had been reduced to about half a dozen proteins and 23S rRNA (for reviews see Ofenand, 1980; Cooperman, 1980), and following the discovery of catalytic RNAs (Guerrier-Takada et al., 1983; Kruger et al., 1982), the hypothesis that 23S rRNA might be its sole constituent, which had been proposed years earlier, began to gain favor. In 1984, Noller and colleagues published affinity labeling results which showed that U2619 and U2620 (in E. coli: U2584, U2585) are adjacent to the CCA-end of P site-bound tRNA (Barta et al., 1984; Vester et al., 1988). These nucleotides are part of a highly conserved internal loop in the center of domain V of 23S rRNA. The hypothesis that this loop is intimately involved in the peptidyl transferase activity was supported by the observation that mutations in that loop render cells resistant to many inhibitors of peptidyl transferase, and evidence implicating it in this activity has continued to mount (see Noller, 1991; Garrett et al., 1996).
Definitive proof that the central loop in domain V is the sole component of the ribosome involved in the peptidyl transferase activity has remained elusive, however. In the 1990s, Noller and his colleagues prepared particles that retain peptidyl transferase activity by increasingly vigorous deproteinizations of large ribosomal subunits, but active particles that were completely protein-free could not be produced (Noller et al., 1999; Khaitovich et al., 1999). Nevertheless, combined with earlier reconstitution results (Franceschi et al., 1990), this work reduced the number of proteins that might be involved to just two: L2 and L3 (see Green et al., 1997). More recently, Watanabe and coworkers reported success in eliciting peptidyl transferase activity from in vitro synthesized, protein-free 23S mRNA (Nitta et al., 1998; Nitta et al., 1998), but their observations have not withstood further scrutiny (Khaitovich et al., 1999). Thus the question still remains: is the ribosome a ribozyme or is it not?
Over the years, the location of the peptidyl transferase site in the ribosome has been approached almost exclusively by electron microscopy. In the mid-1980s evidence that there is a tunnel running through the large ribosomal subunit from the middle of its subunit interface side to its back (Milligan et al., 1986; Yonath et al., 1987) began to accumulate, and there was/is strong reason to believe that polypeptides pass through it as they are synthesized (Bernabeu et al., 1982; Ryabova et al., 1988; Beckmann et al., 1997). More recent cryo-electron microscopic investigations (Frank et al., 1995; Frank et al., 1995; Stark et al., 1997; Stark et al., 1995) confirmed the existence of the tunnel and demonstrated that the CCA-ends of ribosome-bound tRNAs bound to the A- and P-sites are found in the subunit interface end of the tunnel. Consequently, the peptidyl transferase site must be located at that same position, which is at the bottom of a deep cleft in the center of the subunit interface surface of the large subunit, immediately below its central protuberance.
The substrates of the reaction catalyzed by the large subunit are an aminoacyl-tRNA (aa-tRNA) and a peptidyl-tRNA. The former bind in the ribosome's A-site and the latter in its P-site. The α-amino group of the aa-tRNA attacks the carbon of the carbonyl acylating the 3′ hydroxyl group of the peptidyl-tRNA, and a tetrahedral intermediate is formed at the carbonyl carbon (FIG. 1). The tetrahedral intermediate resolves to yield a peptide extended by one amino acid esterified to the A-site bound tRNA and a deacylated tRNA in the P-site.
This reaction scheme is supported by the observations of Yarus and colleagues (Welch et al., 1995) who synthesized an analogue of the tetrahedral intermediate by joining an oligonucleotide having the sequence CCdA to puromycin via a phosphoramide group (FIG. 9). The sequence CCA, which is the 3′ terminal sequence of all tRNAs, binds to the large subunit by itself, consistent with the biochemical data showing that the interactions between tRNAs and the large subunit largely depend on their CCA sequences (Moazed et al., 1991). Puromycin is an aa-tRNA analogue that interacts with the ribosomal A-site, and the phosphoramide group of their compound mimics the tetrahedral carbon intermediate. This transition state analogue, CCdA-phosphate-puromycin (CCdA-p-Puro), binds tightly to the ribosome, and inhibits its peptidyl transferase activity (Welch et al., 1995).
Structure Determination of Macromolecules using X-Ray Crystallography
Each atom in a crystal scatters X-rays in all directions, but crystalline diffraction is observed only when the crystal is oriented relative to the X-ray beam so that the atomic scattering interferes constructively. The orientations that lead to diffraction may be computed if the wavelength of the X-rays used and the symmetry and dimensions of the crystal's unit cell are known (Blundell, T. L. and Johnson, L. N., Protein Crystallography, 1976, Academic Press, New York). The result is that if a film is placed behind a crystal that is being irradiated with monochromatic X-rays of an appropriate wavelength, the diffraction pattern recorded will consist of spots, each spot representing one of the orientations that gives rise to constructive interference.
Each spot in such a pattern, however it is recorded, is characterized by an intensity—its blackness—, a location, which encodes the information about diffraction orientation, and a phase. If all of those things are known about each spot in a crystal diffraction pattern, the distribution of electrons in the unit cell of the crystal may be computed by Fourier transformation (Blundell, T. L. and Johnson, L. N., supra), and from that distribution or electron density map, atomic positions can be worked out.
Unfortunately, the phase information essential for computing electron distributions cannot be measured directly from diffraction patterns. One of the methods routinely used to determine the phases of macromolecules, such as proteins and nucleic acids, is called multiple isomorphous replacement (MIR) (including heavy metal scattering), which requires the introduction of new x-ray scatterers into the unit cell of the crystal. These additions are usually heavy atoms (so that they make a significant contribution to the diffraction pattern), such that there should not be too many of them (so that their positions can be located); and they should not change the structure of the molecule or of the crystal cell, i.e., the crystals should be isomorphous. Isomorphous replacement is usually done by diffusing different heavy-metal complexes into the channels of the preformed protein crystals. The protein molecules expose side chains (such as SH groups) into these solvent channels that are able to bind heavy metals. It is also possible to replace endogenous light metals in metalloproteins with heavier ones, e.g., zinc by mercury, or calcium by samarium. Alternatively, the isomorphous derivative can be obtained by covalently attaching a heavy metal to the macromolecular in solution and then to subject it to crystallization conditions.
Heavy metal atoms routinely used for isomorphous replacement include but are not limited to mercury, uranium, platinum, gold, lead, and selenium. Specific examples include mercury chloride, ethyl-mercury phosphate, and osmium pentamine, iridium pentamine.
Since such heavy metals contain many more electrons than the light atoms (H, N, C, O, and S) of the protein, they scatter x-rays more strongly. All diffracted beams would therefore increase in intensity after heavy-metal substitution if all interference were positive. In fact, however, some interference is negative; consequently, following heavy-metal substitution, some spots measurably increase in intensity, others decrease, and many show no detectable difference.
Phase differences between diffracted spots can be determined from intensity changes following heavy-metal substitution. First, the intensity differences are used to deduce the positions of the heavy atoms in the crystal unit cell. Fourier summations of these intensity differences give maps of the vectors between the heavy atoms, the so-called Patterson maps. From these vector maps the atomic arrangement of the heavy atoms is deduced. From the positions of the heavy metals in the unit cell, one can calculate the amplitudes and phases of their contribution to the diffracted beams of protein crystals containing heavy metals.
This knowledge is then used to find the phase of the contribution from the protein in the absence of the heavy-metal atoms. As both the phase and amplitude of the heavy metals and the amplitude of the protein alone is known, as well as the amplitude of the protein plus heavy metals (i.e., protein heavy-metal complex), one phase and three amplitudes are known. From this, the interference of the X-rays scattered by the heavy metals and protein can be calculated to see if it is constructive or destructive. The extent of positive or negative interference, with knowledge of the phase of the heavy metal, given an estimate of the phase of the protein. Because two different phase angles are determined and are equally good solutions, a second heavy-metal complex can be used which also gives two possible phase angles. Only one of these will have the same value as one of the two previous phase angles; it therefore represents the correct phase angle. In practice, more than two different heavy-metal complexes are usually made in order to give a reasonably good phase determination for all reflections. Each individual phase estimate contains experimental errors arising from errors in the measured amplitudes. Furthermore, for many reflections, the intensity differences are too small to measure after one particular isomorphous replacement, and others can be tried.
The amplitudes and the phases of the diffraction data from the protein crystals are used to calculate an electron-density map of the repeating unit of the crystal. This map then has to be interpreted as a polypeptide chain with a particular amino acid sequence. The interpretation of the electron-density map is made more complex by several limitations of the data. First of all, the map itself contains errors, mainly due to errors in the phase angles. In addition, the quality of the map depends on the resolution of the diffraction data, which in turn depends on how well-ordered the crystals are. This directly influences the image that can be produced. The resolution is measured in Å units; the smaller this number is, the higher the resolution and therefore the greater the amount of detail that can be seen.
Building the initial model is a trial-and-error process. First, one has to decide how the polypeptide chain weaves its way through the electron-density map. The resulting chain trace constitutes a hypothesis, by which one tries to match the density of the side chains to the known sequence of the polypeptide. When a reasonable chain trace has finally been obtained, an initial model is built to give the best fit of the atoms to the electron density. Computer graphics are used both for chain tracing and for model building to present the data and manipulated the models.
The initial model will contain some errors. Provided the protein crystals diffract to high enough resolution (e.g., better than 3.5 Å), most or substantially all of the errors can be removed by crystallographic refinement of the model using computer algorithms. In this process, the model is changed to minimize the difference between the experimentally observed diffraction amplitudes and those calculated for a hypothetical crystal containing the model (instead of the real molecule). This difference is expressed as an R factor (residual disagreement) which is 0.0 for exact agreement and about 0.59 for total disagreement.
In general, the R factor is preferably between 0.15 and 0.35 (such as less than about 0.24-0.28) for a well-determined protein structure. The residual difference is a consequence of errors and imperfections in the data. These derive from various sources, including slight variations in the conformation of the protein molecules, as well as inaccurate corrections both for the presence of solvent and for differences in the orientation of the microcrystals from which the crystal is built. This means that the final model represents an average of molecules that are slightly different both in conformation and orientation.
In refined structures at high resolution, there are usually no major errors in the orientation of individual residues, and the estimated errors in atomic positions are usually around 0.1-0.2 Å, provided the amino acid sequence is known. Hydrogen bonds, both within the protein and to bound ligands, can be identified with a high degree of confidence.
Most x-ray structures are determined to a resolution between 1.7 Å and 3.5 Å. Electron-density maps with this resolution range are preferably interpreted by fitting the known amino acid sequences into regions of electron density in which individual atoms are not resolved.
In summary, determining the structure of a macromolecule employs, but are not limited to the following steps (Cantor, C. R. and Schimmel, P. R., Biophysical Chemistry, Part II, Chapter 13, pages 763-764, 1980, Freeman and Company, San Francisco):
1. Prepare crystals of the native macromolecules. Using the crystals, determine the space group and unit-cell dimensions, and collect a set of scattering amplitude data.
2. Prepare heavy-atom isomorphous derivatives. For each derivative, collect a new set of scattering amplitude data.
3. Find the locations of the heavy atoms in the crystal. A routinely practiced method is the difference isomorphous Patterson synthesis.
4. Refine the positions assigned to heavy atoms using difference Fourier refinement techniques or other methods that are known to crystallographers.
5. Estimate the phases of the parent crystal(s) by comparing the structure factor data of the parent crystal with the corresponding data of one or more heavy-atom isomorphous derivatives. Generally, the more heavy-atom derivatives available, the more accurate the phase estimates will be.
6. Refine the positions of the heavy atoms further using least-square, difference Fourier techniques, or other methods known in the art.
7. Calculate an electron density map using the estimated phase and observed amplitude information.
8. Build a model based on the electron density map. Repeating steps 4-8 with data at higher resolution to construct a molecular model.
One can also attempt to refine the structure by calculating phases from the atom positions in the molecular model and use them in stead of the phases determined in steps 5 and 6. The refinement can involve Fourier transform or least-squares techniques, and can treat just the x-ray data or can also include information about known energetics of protein conformation.